Core circadian clock genes control molecular and behavioral circatidal rhythms in Parhyale hawaiensis

Victoria Louis; Zachary Bellido; Adam Helfenbein; Joshua J C Rosenthal; Patrick Emery

doi:10.1016/j.cub.2026.08.017

. Author manuscript; available in PMC: 2026 Oct 2.

Published in final edited form as: Curr Biol. 2026 Sep 2;36(18):4749–4764.e6. doi: 10.1016/j.cub.2026.08.017

Core circadian clock genes control molecular and behavioral circatidal rhythms in Parhyale hawaiensis

Victoria Louis ¹, Zachary Bellido ², Adam Helfenbein ^1,³, Joshua J C Rosenthal ^2,^*, Patrick Emery ^1,^4,^*

PMCID: PMC13629695 NIHMSID: NIHMS2208805 PMID: 42685692

Summary

Marine organisms exhibit 12.4-hour rhythms of gene expression, physiology and behavior synchronized by tidal cues. The mechanism underlying these circatidal rhythms, and its overlap with the circadian clockwork, has remained elusive. However, recent studies showed that the core circadian gene BMAL1 sustains circatidal behavior in crustaceans. Therefore, we mutagenized the other three core circadian clock genes (PhCry2, PhPer and PhClk) in P. hawaiensis, a marine amphipod. We found that they are necessary for both circadian and circatidal behaviors. Moreover, all four core circadian genes are critical for 24-h oscillations of mRNA levels in circadian brain neurons and 12.4-h mRNA rhythms in circatidal neurons. Unexpectedly, the mutants indicate that PhCLK represses PhPer expression independently of PhBMAL1 specifically in circatidal neurons. Our study thus reveals that circadian and circatidal clocks share four core molecular components, but their transcriptional wiring differs.

Graphical Abstract

graphic file with name nihms-2208805-f0001.webp

Etoc blurb:

The12.4-h circatidal clock allows marine animals to cope with the tides. Louis et al. show that in the crustacean P. hawaiensis, this timekeeper is comprised of the same four core proteins as the circadian clock, but that the transcriptional wiring between these components differs in circadian and circatidal brain neurons.

Introduction

Organisms adjust their metabolism, physiology and behavior to predictable cycles in their environment using biological clocks^1–4. Among them, circadian clocks are best understood. They entrain to the alternance of day and night and free-run with a ~24-hour (h) period under constant conditions. The circadian clock mechanism is highly conserved across animals^1,5. Its core is a transcriptional feedback loop comprised of the activators CLOCK (CLK) and BMAL1 that heterodimerize to bind to E-boxes and activate the transcription of numerous genes, including those encoding their own repressors PERIOD (PER), CRYPTOCRHOME (CRY) and/or TIMELESS (TIM), depending on the species. Other CLK/BMAL1 targets include transcription factors that form interlocked loops that modulate and strengthen the core loop^5–11.

In marine environments, organisms are subjected to environmental changes caused by tides. In response, besides circadian rhythms, they exhibit ~12.4-h rhythms that synchronize to tidal cues^12,13. The molecular mechanism behind circatidal rhythms remains poorly characterized. Early behavioral studies in intertidal organisms led to three main hypotheses. The first posited that circadian and circatidal rhythms are generated by distinct clocks¹⁴. The second proposed that circadian and circatidal rhythms are produced by a single clock, which can run with a period of ~24 h or ~12.4 h, depending on the rhythmic cues an animal is exposed to¹⁵. The last hypothesis suggested the existence of two circalunidian (~24.8-h) clocks running in antiphase, thus generating circatidal outputs¹⁶. This third hypothesis could involve essentially the same mechanism as the circadian clock, with a minor adjustment of the oscillator’s period (from 24 to 24.8 h). A similar period adjustment could permit the circadian clock to generate ~12.4-h rhythms by exploiting its two antiphasic interlocked transcriptional feedback loops, or by driving neuronal activity in opposite phases in two distinct subsets of pacemaker neurons, as observed for circadian morning and evening neurons in Drosophila^17,18.

Several studies have tested whether circadian clock genes are required for circatidal rhythms. Knock downs via RNA interference (RNAi) in the mangrove cricket Apteronemobius asahinai and the isopod Eurydice pulchra suggested that Clk, Per and Cry2 are not required to sustain circatidal outputs^19–22. However, pharmacological inhibition of Casein Kinase Iε, which phosphorylates PER proteins and thus regulates circadian rhythms in flies and mammals²³, lengthened circatidal behavioral rhythms, suggesting that this kinase is shared between the circadian and circatidal clocks²¹. Recent studies further pointed towards a mechanistic overlap between the two clocks. CRISPR-Cas9 mediated knock out of PhBmal1 in the amphipod Parhyale hawaiensis eliminated circatidal behavior²⁴, and RNAi directed at Bmal1 reduced circatidal rhythmicity in E. pulchra²². Furthermore, distinct circadian and circatidal clocks cells were identified in the brain of P. hawaiensis and E. pulchra²⁵. In P. hawaiensis, ~12.4-h rhythms of expression for both PhPer and PhCry2 mRNAs were observed in four out of the ~60 cells expressing core circadian genes. These oscillations were synchronized by a tidal cue (i.e. vibration), but not by the light/dark (LD) cycle. By contrast, circadian cells displayed 24-h period rhythms of PhPer and PhCry2 mRNAs that synchronized to the LD cycle, but not to vibration. These data suggest that PhPer and PhCry2 contribute to circatidal rhythms in P. hawaiensis.

Here, we used CRISPR-Cas9 genome editing to test the degree of mechanistic overlap between the circadian and circatidal clocks in P. hawaiensis by generating knock-out lines for PhPer, PhClk and PhCry2 to complement the existing PhBmal1 knock-out animals. Interestingly, we found that all four core clock genes are necessary to sustain circatidal rhythms, indicating considerable mechanistic overlap between the two clock mechanisms. Nonetheless, our results also suggest that the transcriptional wiring between the four core clock genes differs between circadian and circatidal cells.

Results

Loss of PhCry2 disrupts circadian behavior

Based on genomic studies, the circadian clock of P. hawaiensis is predicted to comprise homologs of CLK, BMAL1, PER and repressor-type cryptochromes²⁶. The genome does not appear to encode a photoreceptive CRY or a TIM homolog. We previously demonstrated that PhBMAL1 is required for both circadian and circatidal behavioral rhythms²⁴. We therefore decided to determine whether the circadian and circatidal clocks share additional common building blocks.

We first targeted the PhCry2 gene. Two knock-out lines were generated using CRISPR-Cas9 and two guide RNAs (gRNA) targeting the photolyase domain (Figure 1A), which is required for cryptochromes’ repressor function²⁷. The first line (PhCry2-mutation 1) harbored two deletions of 18 and 20 base pairs (bp), respectively, which cause a premature stop codon that truncates PhCRY2 from 929 amino acids (AA) to 422 AA. The second line (PhCry2-mutation 2) harbored a single deletion of 16 bp, also causing a premature termination codon that truncates PhCRY2 to 428 amino acids.

To verify that the truncation affects PhCRY2 activity, we turned to a transcriptional assay in mammalian HEK-293T cells (Figure 1B). We previously showed that PhBMAL1 can activate E-box mediated transcription with mCLK²⁴ (there is still uncertainty on the exact sequence of PhCLK, see below). We tested if PhPER and PhCRY2 repress PhBMAL1/mCLK, as in other animals^21,28–32. Indeed, expression of both PhPER and PhCRY2 reduced PhBMAL1/mCLK activity in a dose-dependent manner. Repression was also observed with PhCRY2 alone but was not as efficient as with PhPER (two-way ANOVA, *p=0.01). PhPER did not repress on its own. Thus, as in other animals, PhCRY2 appears to be the primary repressor, while PhPER plays a modulatory function^{21,29,30,32,33}. No repression was observed with the truncated PhCRY2 - despite comparable levels of expression to wild-type (Figures 1B, S1A and S1B). PhCRY2- is thus severely defective.

To assess whether the loss of PhCRY2 disrupts the circadian clock, we measured the swimming activity of mutant and control animals under constant conditions after entrainment to a LD cycle. We prioritized free-running behavior under constant light (LL), as we previously observed that circadian behavioral rhythms are more robust under these conditions than under constant darkness (DD)²⁴. This was also the case in the present study (Figures 1C and S2A), although rhythms were weaker and more likely to show a bimodal daily pattern of activity than previously reported under both conditions (see “behavior analysis” in method section for more detail, and Table S1). As expected, under LL, average activity traces of wild-type (WT) animals (PhCry2 +/+) showed lower activity during the subjective light phase and higher activity during the subjective dark phase (Figures 1C and S1C). The heterozygous animals (PhCry2 +/−) exhibited a similar pattern. In contrast, the average swimming activity of the homozygous mutants (PhCry2 −/−) showed no obvious rhythmicity. This was reflected in the proportion of rhythmic animals (Figure 1D) and in the rhythm power (amplitude), which tended to be weaker in the rare rhythmic knockout animals (Figure 1E). We next determined the period of behavioral rhythms using periodogram analysis. After LD-only entrainment, most rhythmic animals exhibit statistically significant rhythm periodicities in the circadian (~24-h) range (“doublet”), but harmonics in the ~12-h range can also be observed (“singlets”)²⁴. The average circadian doublet period of WT animals was around 25.5 h because continuous light exposure lengthens circadian rhythms in P. hawaiensis²⁴ (Figure S3E and Table S2). While the average circadian period did not statistically differ between wild-type, heterozygous and the rare rhythmic homozygous mutants, individual periodicities were not clustered around 25.5 h in mutant animals, as in wild-type and heterozygous animals (Figure 1F). Out of the four rhythmic PhCry2 −/−, only one had a period within the expected range. The singlet period of the homozygous mutants was also very variable and significantly longer than WT (Figure 1G).

To further establish that PhCRY2 controls circadian rhythms, we tested LD-only entrained animals in DD. Again, circadian behavior was severely disrupted (Figures S2A and B). Only two knockout animals out of 32 were rhythmic and only one of them had a period within the expected range (Figures 1H to 1K). Therefore, our results show that in the absence of functional PhCRY2, circadian behavior is severely compromised. As previously observed with PhBmal1−/− mutants, residual low amplitude rhythms with poorly defined period emerge in a few animals²⁴.

Loss of PhCry2 disrupts circadian molecular rhythms

In Drosophila, ca. 240 neurons rhythmically express circadian clock genes³⁴. These neurons drive different 24-h metabolic and behavioral rhythms. In addition, numerous glial cells express core clock genes^35–37. In the P. hawaiensis brain, only ~ 60 cells detectably express PhPer and PhCry2²⁵. To assess the effect of the PhCry2 knock-out on the molecular circadian clock, we turned to HCR-FISH to monitor PhPer and PhCry2 mRNA rhythms in putative circadian neurons, the medioposterior (MP) cells²⁵. These cells were found to entrain to the LD cycle, showing 24-h mRNA abundance rhythms after entrainment to both a 24-h LD cycle and a 12.4-h vibration cycle that mimics tides²⁵. Using probes to the pan-neuronal marker elav, we established that most MP cells indeed are neurons, although 1/10 cell per hemisphere appeared elav-negative (Figure S3A). Since we use immersion cycles instead of vibration for tidal entrainment, we verified that the MP neurons exhibit circadian mRNA rhythms in our hands. Indeed, we observed circadian PhCry2 and PhPer expression in constant condition after entrainment to combined LD and tides in WT (Figures S4A and B). As surprisingly observed by Oliphant et al., PhCry2 and PhPer mRNAs also oscillated with an opposite phase after exposure to our entrainment conditions. While the PhPer expression pattern was clearly circadian, PhCry2 showed a statistically significant 12-h period rhythm according to JTK cycle analysis, although the secondary peak was separated from the main peak by about 8 h. This is consistent with Oliphant et al., who could also observe a secondary PhCry2 mRNA peak when varying the phase relationship between the LD and vibration cycles²⁵. In PhCry2 −/− animals, PhCry2 mRNA levels remained constantly low (Figures 1L and M). PhPer expression was also significantly disrupted as mRNA levels were significantly higher than in WT, except at CT18. A weak, but significant decrease in PhPer2 mRNA level was observed at CT12 and 18, but the phase of this weak apparent rhythm was completely distinct from that of the robust rhythms observed in WT. A similar progressive decrease in mPer1-2 mRNA level has been observed in mCry1 −/− mCry2 −/− double-mutant mice, perhaps as a result of prior mPer1 and mPer2 photic induction³⁸. The slight decrease in PhPer mRNA over time might thus be the result of prior light exposure. Taken together, our results show that PhCRY2 is part of the molecular circadian clock and necessary for circadian behavior.

PhCry2 is an essential element of the circatidal clock

Next, we determined the role of PhCRY2 in the circatidal clock. We entrained animals to combined LD and tidal (10.3 h high tide: 2.1 h low tide) cycles and monitored their behavior under DD and constant high tide (HT) conditions. As expected²⁴, WT and heterozygous animals showed strong activity rhythms (Figures 2A and S2D). Low activity was observed at subjective low tide and activity peaked during the subjective high tide. In contrast, the homozygous mutants did not show any rhythms of activity. All PhCry2 −/− animal scored as arrhythmic (Figure 2B). A slight gene dosage effect was observed in heterozygous animals compared to WT, as the period of their circatidal behavior was significantly longer (Figures 2C, D and E). These results show that PhCRY2 is necessary to generate circatidal behavior.

Figure 2. — A) Average relative swimming activity of wild-type (*PhCry2*+/+), heterozygous (*PhCry2*+/−) and homozygous mutant (*PhCry2*−/−) animals recorded in DD and high tide after entrainment to a LD and tidal (10.3 h high tide: 2.1 h low tide) cycles. Blue shading represents the expected high tide. Standard error to the mean (SEM) is represented with the orange shading.

B) Percentage of rhythmic animals under DD and high tide (Chi-square test, ***p <0.001; subsequent Chi-square test, +/+ vs +/−, p = 0.86, +/+ vs −/−, ***p <0.001 and +/− vs −/−, ***p <0.001).

C) Rhythm power averages under DD and high tide (Welch’s t test, p = 0.416).

D) Singlet period averages under DD and high tide (Welch’s t test, **p = 0.007).

E) Doublet period averages under DD and high tide (Welch’s t test, **p = 0.005).

F) HCR-FISH of *PhPer* and *PhCry2* mRNAs in circatidal dorsal-lateral clock neurons exposed to constant conditions after entrainment to 10.3:2.1 tides and 12:12 LD cycles in *PhCry2*−/− animals (*PhCry2* – mutation 1), as represented in Figure 1.

G) As represented in Figure 1, quantification of the number of HCR-FISH spots in *PhCry2* −/− animals. In the mutant, there was an effect of time on gene expression for *PhPer*, (Kruskal-Wallis, *p = 0.0159), but not for *PhCry2* (Kruskal-Wallis, p = 0.545) (See Table S5 for post-hoc test p-value). Statistical differences between genotypes are indicated with * signs (Tukey’s multiple comparison, See Table S6 for exact p-value).

See also Figure S2–S4, Table S1, S2, S5, S6, S7, and Data S1D.

To assess the implication of PhCry2 in molecular circatidal rhythms, we returned to HCR-FISH. The expression of PhCry2 and PhPer was this time measured in the dorsal-lateral (DL) cells, which we found to be neurons as well (Figure S3B). The DL neurons entrain to vibration, exhibit ca. 12-h mRNA rhythms and are thus candidate circatidal oscillators²⁵. In WT, both genes showed robust 12-h rhythms of expression with low levels around the expected low tide (Figures S4C and S4D). In PhCry2 −/−, as observed in circadian neurons, PhCry2 mRNA levels remained constantly low (Figures 2F and G). Also, similarly to circadian neurons, PhPer expression was high and decreased over time, but did not show any sign of residual circatidal expression. In summary, our behavioral and molecular results demonstrate that PhCRY2 is an essential circatidal protein and is thus shared between the circadian and the circatidal systems.

Loss of PhPer disrupts circadian rhythms

PhPER contains two Per-ARNt-Sim (PAS) domains and a Per-ARNt-Carboxy-terminal (PAC) domain. Three knock-out mutant lines were generated using CRISPR-Cas9 and two gRNAs targeting an exon preceding those encoding the key functional domains (Figure 3A). The first line (PhPer – mutation 1) harbored a 5’ 8 bp deletion combined with 2 added bp and a 3’ 25-bp deletion with 4 added bp. These indels thus cause a premature stop codon truncating PER from 1258 to 333 AA, eliminating all key functional domains. The second line (PhPer – mutation 2) had a premature stop codon following a 5’ deletion of 10 bp combined with the addition of 5 bp and a 3’ deletion of 2 bp, thus truncating PhPER to 317 AA. The last line (PhPer – mutation 3), similarly to the others, had a premature stop codon and a truncated protein of 355 AA after a 5’ deletion of 9 bp with 3 extra bp and a 3’ deletion of 7 bp. The absence of functional domains is predictive of a complete loss of function for all three mutations. Indeed, truncated PhPER did not facilitate PhCRY2 repression of mCLK/PhBMAL1 in HEK293 cells (Figures S1C and D).

Figure 3. — A) Schematics of *PhPer* mutagenesis. Blue and red boxes indicate the gene sequences targeted by the gRNAs used for CRISPR/Cas9 mutagenesis. Three knock-out mutant lines with premature stop codons were obtained. The resulting mutant PhPER proteins are very similar, with slightly different C-terminal tails (see boxed amino acid sequences). All lines are missing key functional domains: the two Per-ARNt-Sim (PAS) (yellow) and the Per-ARNt-Carboxy-terminal (PAC) domains (blue). Thus, behavior data from the three strains were combined on this figure and on Figure 4.

B) Average relative swimming activity of wild-type (*PhPer*+/+), heterozygous (*PhPer*+/−) and homozygous mutant (*PhPer*−/−) animals recorded in constant light (LL) after entrainment to a LD cycle. Grey shading represents the subjective night. Standard error to the mean (SEM) is represented with the orange shading.

C) Percentage of rhythmic animals in LL (Chi-square test, *p = 0.049; subsequent Chi-square test, +/+ vs +/−, p = 0.93, +/+ vs −/−, *p = 0.023 and +/− vs −/−, *p = 0.016).

D) Rhythm power averages under LL (Kruskal-Wallis test, p = 0.137).

E) Doublet period averages under LL (Kruskal-Wallis test, p = 0.544).

F) Singlet period averages under LL (Welch’s test, p = 0.938).

G) Percentage of rhythmic animals under DD after LD entrainment (Chi-square test, ***p < 0.001; subsequent pairwise Chi-square test, ***p < 0.001).

H) Rhythm power averages under DD (Mann-Whitney Smirnov test, p = 0.492).

I) Doublet period averages under DD (Welch’s test, *p = 0.043).

J) Singlet period averages under DD (Welch’s test, *p = 0.045).

K) HCR-FISH to *PhPer* and *PhCry2* mRNAs in circadian medioposterior clock neurons exposed to constant conditions after entrainment to 10.3:2.1 tidal and 12:12 LD cycles in *PhPer* −/− animals (*PhPer* – mutation 1 and 3) as represented in Figure 1.

L) As represented in Figure 1, quantification of the number of HCR-FISH spots in *PhPer* −/− animals. After testing the effect of time on the gene expression (*PhPer*, Kruskal-Wallis, ***p < 0.001, *PhCry2*, Kruskal-Wallis, ***p < 0.001), rhythm was tested using the JTK-cycle algorithm for 24-h periodicity (See Table S3 for exact p-value). Statistical differences between genotypes are indicated with * signs (Tukey’s multiple comparison, See Table S4 for exact p-value).

See also Figures S1, S3–S5, Table S1–S4, S7, Data S1C, E and F.

Again, we first measured circadian behavior in LL. Both WT (PhPer +/+) and heterozygous animals (PhPer +/−) showed rhythmic behavior at the population level (Figures 3B and S5A). The homozygous mutant (PhPer −/−) population showed no clear rhythmic activity pattern. Accordingly, the percentage of rhythmic mutant animals was significantly lower than in PhPer +/+ and PhPer +/− populations (Figure 3C). Rhythm power of mutant animals was very low, but significance was not reached compared to wild-type and heterozygous, since only two animals were rhythmic and statistical power was thus low (Figure 3D). Out of the two rhythmic PhPer −/− animals, only one had a period within the expected range (Figures 3E and F). In DD, homozygous mutants were significantly less rhythmic than WT with only four rhythmic animals out of 72 (Figures 3G to J and S5D and S5E). These results show that PhPer is critical to sustain circadian behavior.

Accordingly, both PhCry2 and PhPer mRNA rhythms were disrupted in the circadian MP neurons. mRNA levels decreased over time instead of showing peaks of expression in the evening or night for PhCry2 and PhPer, respectively (Figures 3K and L), reminiscent of the decay observed for PhPer mRNA in PhCry2 −/− animals. Altogether, these data show that loss of PhPER disrupts both molecular and behavioral circadian rhythms.

PhPer sustains circatidal behavior as part of the circatidal clock

We then determined the role of PhPer in the circatidal clock. After entrainment to combined LD cycle and tides (last high tide [LHT] 13:20), both WT and heterozygous animals showed lower average activity around the subjective low tides and higher activity around subjective high tides (Figures 4A and S5B). The average activity of animals lacking PhPer also showed this rhythmic pattern for 3 cycles, but it was much noisier and rhythm amplitude decreased rapidly. To determine if this residual rhythmicity was circatidal in nature, animals were entrained to a 6h-shifted tidal cycle (LHT 19:20) (Figures 4B and S5C). Again, PhPer mutants showed lower activity around the subjective low tides for 2, perhaps 3 cycles. The phase of these residual rhythms was shifted by 6 hours, as in WT animals, confirming that they are entrained by tides (Figure 4C). However, the proportion of animals scoring as rhythmic was much lower in PhPer −/− mutants compared to WT and heterozygous animals (Figure 4D). For the few rhythmic mutant animals, power and period did not significantly differ from WT and heterozygotes (Figures 4E, F and G). These results show that PhPER sustains circatidal behavioral rhythms, although it might not be as critical as PhCRY2.

Figure 4. — A) Average relative swimming activity of wild-type (*PhPer*+/+), heterozygous (*PhPer2*+/−) and homozygous mutant (*PhPer*−/−) recorded in constant dark (DD) after entrainment to 10.3:2.1 tidal and 12:12 LD cycles as represented in Figure 2. The last high tide occurred at 13:20.

B) Average relative swimming activity as represented in A. The tidal cycle was shifted by 6 hours; therefore the last high tide occurred at 19:20.

C) Phase of maximum activity in *PhPer*+/+ and *PhPer*−/− under the two tidal regimes tested (One-way ANOVA, ***p <0.001 subsequent Sidak’s multiple comparison, LHT13:20 peak1, p = 0.380, LHT13:20 peak2, p = 0.347, LHT19:20 peak1, p = 0.999 and LHT19:20 peak2, p = 0.945). Within tidal regimes, peak phases are significantly different from each other (LHT13:20, Welch’s test, ***p <0.001 and LHT19:20, Welch’s test, ***p <0.001).

D) Percentage of rhythmic animals under DD and high tide. Tidal regimes were combined (Chi-square test, ***p <0.001; subsequent Chi-square test, +/+ vs +/−, p = 0.50, +/+ vs −/−, ***p <0.001 and +/− vs −/−, ***p <0.001).

E) Rhythm power averages under DD and high tide (Kruskal-Wallis, p = 0.173)

F) Singlet period averages under DD and high tide (Kruskal-Wallis, p = 0.300)

G) Doublet period averages under DD and high tide (Kruskal-Wallis, p = 0.0602)

H) HCR-FISH of *PhPer* and *PhCry2* mRNAs in circatidal dorsal-lateral clock neurons exposed to constant conditions after entrainment to 10.3:2.1 tides and 12:12 LD cycles in *PhPer* −/− animals (*PhPer* – mutation 1 and 3) as represented in Figure1.

I) As represented in Figure 1, quantification of HCR-FISH spots per hemisphere across CT in *PhPer*−/− animals. (*PhPer*, Kruskal-Wallis, p = 0.174, *PhCry2*, Kruskal-Wallis, p=0.056). Statistical differences between genotypes are indicated with * signs (Tukey’s multiple comparison, See Table S6 for exact p-value).

See also Figure S3–S5, Table S1, S2, S5, S6, S7, Data S1G and H.

We then determined the impact of the loss of PhPER on molecular circatidal rhythms in DL neurons. Neither PhPer nor PhCry2 mRNAs showed statistically significant time-dependent variations in abundance (Figures 4H and I). Circatidal molecular rhythms were therefore clearly disrupted. However, we noticed slightly higher levels of both mRNAs at the time at which expression is high in WT animals. Weak residual molecular rhythms might thus be present in PhPer−/− animals, which could explain the observed weak circatidal behavior rhythms. Overall, however, our data show that PhPer is part of the circatidal clock mechanism, although PhPer mutant animals can maintain weak circatidal behavioral rhythms in its absence, contrary to PhCry2 −/− animals.

PhClk is also required for circadian rhythms

CLK is composed of a basic-Helix-Loop-Helix (bHLH) domain that binds E-Boxes and two PAS domains needed for dimerization with BMAL1 (Figure 5A). While a previous study failed to identify CLK’s PAS-A coding sequences in the P. hawaiensis genome or transcriptome²⁶, we found two putative exons containing PAS-A coding sequences (Figure S6), indicating that PhCLK contains all three canonical domains. However, in the absence of transcriptomics support, we still cannot predict the full PhCLK sequence. To knock-out PhClk, two gRNAs were designed, targeting the bHLH domain. Two mutant lines were generated, lacking all functional domains due to an early stop-codon in the bHLH coding region. The first line (PhClk – mutation 1) had a deletion of 27 bp with 4 extra bp inserted, generating a frameshift. The predicted truncated protein was 61 AA long instead of 2065 AA. The second line (PhClk – mutation 2) had a deletion of 80 nucleotides and truncated PhCLK to 42 AA.

Both WT and heterozygous animals showed the expected circadian behavior pattern after LD-only entrainment (Figures 5B and S7A). By contrast, PhClk −/− animals did not show a rhythmic behavior pattern at the population level. Also, we observed fewer rhythmic animals in PhClk −/− compared to WT and heterozygous animals, although the difference did not reach significance (Figure 5C). Similarly, rhythm power was lower in PhClk −/− animals, though not significantly so (Figure 5D). Importantly, periods in the few weakly rhythmic animals were broadly distributed rather than clustered around 25.5 h, as previously observed in weakly rhythmic PhBmal1 mutants²⁴ (Figures 5E and F). Circadian behavior thus appears to be disrupted. Moreover, PhClk was clearly required for circadian behavior under DD, as none of the knock-out animals were rhythmic under these conditions (Figures 5G to J and S7D and E). Furthermore, HCR-FISH revealed that the molecular circadian clock is severely disrupted in the MP neurons of PhClk mutants. Neither PhPer nor PhCry2 mRNA levels oscillated (Figures 5K and L), with extremely low expression level of PhCry2. In summary, PhClk is required for the functioning of the circadian clock in P. hawaiensis and ensures proper circadian swimming activity.

PhClk sustains circatidal behavior as part of the circatidal clock

We then probed the impact of loss of PhCLK on circatidal rhythms at the behavioral and molecular levels. In constant conditions following entrainment to combined LD cycles and tides, both PhClk +/+ and PhClk +/− showed robust rhythmic behavior (Figures 6A and B and S7B and C). Animals lacking PhClk displayed a weakly rhythmic pattern for about 2 circatidal cycles that appeared entrained to tides, reminiscent of our observations with PhPer mutants. The proportion of rhythmic animals and the power of the rhythm were significantly lower in PhClk −/− compared to the other genotypes (Figures 6C and D). No significant difference was observed between genotypes for both the singlet and the doublet periods (Figures 6E and F). Behavior phase was only quantified for the LHT 13:20 as only one PhClk −/− individual was rhythmic with the shifted tidal cycle (Figure 6G). No significant difference of phase was observed between mutants and WT, suggesting tidal entrainment of the weak residual rhythms. These results indicate that PhClk is necessary to sustain circatidal swimming activity.

Figure 6. — A) Average relative swimming activity of wild-type (*PhClk*+/+), heterozygous (*PhClk*+/−) and homozygous mutant (*PhClk*−/−) animals recorded in constant dark (DD) and high tide after entrainment to a LD cycle and tidal (10.3 h high tide: 2.1 h low tide) as represented in Figure 2. The last high tide occurred at 13:20.

B) Relative average swimming as represented in A. The tidal cycle was shifted by 6 hours, so the last high tide occurred at 19:20.

C) Percentage of rhythmic animals under DD and high tide. Tidal regimes were combined (Chi-square test, ***p <0.001; subsequent Chi-square test, +/+ vs +/−, p = 0.564, +/+ vs −/−, ***p <0.001 and +/− vs −/−, ***p <0.001).

D) Rhythm power averages under DD and high tide (Kruskal-Wallis, **p = 0.002; subsequent Dunn’s multiple comparison, +/+ vs +/−, p = 1, +/+ vs −/−, **p = 0.001 and +/− vs −/−, **p = 0.005).

E) Singlet period averages under DD and high tide (Kruskal-Wallis, p = 0.673).

F) Doublet period averages under DD and high tide (Kruskal-Wallis, p = 0.350).

G) Phase of maximum activity in *PhClk*+/+ and *PhClk*−/− in the tidal regime tested (One-way ANOVA, ***p <0.001 subsequent Sidak’s multiple comparison, LHT13:20 peak1, p = 0.982, LHT13:20 peak2, p = 0.946). Within tidal regime, peak phases are significantly different from each other (Welch’s test, ***p <0.001).

H) HCR-FISH to *PhPer* and *PhCry2* mRNAs in circatidal dorsal-lateral neurons exposed to constant conditions after entrainment to 10.3:2.1 tidal and 12:12 LD cycles in *PhClk −*/− animals (*PhClk* – mutation 1) as represented in Figure1.

I) As represented in Figure 1, quantification of HCR-FISH spots in *PhClk*−/− animals (*PhPer*, One-way ANOVA, p =0.370, *PhCry2*, Kruskal-Wallis, p =0.380). Statistical differences between genotypes are indicated with * signs (Tukey’s multiple comparison, See Table S6 for exact p-value).

See also Figures S3, S4, S7, Table S1, S2, S5, S6, S7, Data S1K and L.

At the molecular level, both PhPer and PhCry2 mRNAs were detected in circatidal DL neurons of PhClk mutants (Figures 6H and I) and showed no time-dependent changes in expression. PhPer mRNA levels were constantly high compared to WT, while PhCry2 expression was constantly low. This shows that PhClk is also required for the functioning of the circatidal molecular clock.

PhBmal1 also controls circadian and circatidal expression, but PhBMAl1 and PhCLK functions differ in circatidal neurons

Our previous work showed that PhBmal1 is required to maintain both circadian and circatidal activity in P. hawaiensis, but its impact on the molecular circadian and circatidal clock was not tested²⁴. We thus performed FISH-HCR on PhBmal1 knock-out animals. In the MP neurons, the circadian expression of PhPer and PhCry2 was disrupted (Figures 7A and B). The expression of PhPer decreased over time whereas PhCry2 expression was low at all time points. Nevertheless, PhCry2 showed a weak rhythm with a completely abnormal phase. We therefore conclude that PhBMAL1 is necessary for proper circadian molecular rhythms. In the DL neurons, neither repressor genes showed circatidal expression (Figures 7C and D). This confirms that BMAL1 is part of both the circadian and the circatidal clocks systems in P. hawaiensis.

Figure 7. — A) HCR-FISH to *PhPer* and *PhCry2* mRNAs in circadian medioposterior clock neurons exposed to constant conditions after entrainment to 10.3:2.1 tidal and 12:12 LD cycles in *PhBmal1* −/− animals as represented in Figure 1.

B) As represented in Figure 1, quantification of the number of HCR-FISH spots in *PhBmal1* −/− animals. After testing the effect of time on the gene expression (*PhPer*, One-way ANOVA, ***p < 0.001, *PhCry2*, One-way ANOVA, **p = 0.002), rhythm was tested using the JTK-cycle algorithm for 24 h (See Table S3 for exact p-value). Statistical differences between genotypes are indicated with * signs (Tukey’s multiple comparison, See Table S4 for exact p-value).

C) HCR-FISH of *PhPer* and *PhCry2* mRNAs in circatidal dorsal-lateral clock neurons exposed to constant conditions after entrainment to 10.3:2.1 tides and 12:12 LD cycles in *PhBmal1* −/− animals as represented in Figure 1.

D) As represented in Figure 1, quantification of HCR-FISH spots in *PhBmal1*−/− animals. (*PhPer*, One-way ANOVA, p =0.376, *PhCry2*, One-way ANOVA, p =0.098). Statistical differences between genotypes are indicated with * signs (Tukey’s multiple comparison, See Table S6 for exact p-value).

E) Circadian and circatidal clock models for *P. hawaiensis*. Activators are in green, and repressors in red. Based on our mutant analysis, we propose that the two clocks are wired differently, transcriptionally. A key difference is PhCLK’s role in *PhPer* regulation. In the circadian clock, PhCLK promotes *PhPer* transcription. However, in the circatidal clock, it is predominantly a *PhPer* repressor. It might do so through direct binding to the *PhPer* promoter, or by promoting the expression of an unknown transcriptional repressor. PhBmal1 promotes *PhPer* expression in circatidal neurons, either with the help of *PhClk*, or a different partner. In the circadian neurons, *PhPer* and *PhCry2* mRNAs cycle in antiphase. This could be because *PhCry2* is indirectly regulated by *PhBmal1* and *PhClk*, through posttranscriptional regulation, or perhaps because PhCLK also directly or indirectly regulates *PhCry2* with an unknown partner in addition to *PhBmal1*. See main text for more discussion.

See also Figure S3, S4 and Table S3–S6.

As expected since BMAL1 is a transcriptional activator, PhPer and PhCry2 mRNAs remained constantly low in both circadian and circatidal neurons, never reaching WT peak expression. This was also the case in brain circadian neurons of PhClk mutants (Figure 5K and L), as predicted since CLK and BMAL1 work as heterodimers. In stark contrast, while PhCry2 mRNA was as expected also low, PhPer mRNA was constantly high in circatidal neurons of PhClk mutants (Figure 6H and I). PhCLK thus represses, rather than activates, PhPer expression specifically in circatidal neurons. This reveals a surprising difference in the transcriptional wiring of core clock genes between circadian and circatidal neurons, and a novel PhBMAL1-independent function for PhCLK (Figure 7E).

Since mutant and wild-type animals were processed and imaged separately (see Methods), we repeated the FISH-HCR experiments at CT6 and CT18, processing wild-type and PhClk mutants together to ensure that the unexpected observation that PhPer mRNA levels are high in circatidal neurons of PhClk mutant was not the result of experimental variations or batch effect (Figure S4F). Results were remarkably similar in this independent experiment: PhPer mRNA was again high in circatidal neurons of PhClk mutants while PhCry2 mRNA was at trough levels. Also, in circadian neurons, PhCRY2 mRNA levels were again essentially undetectable.

Discussion

We previously showed that PhBmal1 is necessary for circatidal behavior in P. hawaiensis, in addition to its canonical role in the circadian clock²⁴. Here, we find that there is considerable overlap between circadian and circatidal clock mechanisms. We demonstrate that the three other core circadian genes are required to sustain circatidal behavior. This is in stark contrast to previous studies in E. pulchra and A. asahinai^19–22. The absence of RNAi phenotypes when targeting Per, Clk and Cry2 in these organisms might have resulted from incomplete suppression of gene expression. In support of this idea, RNAi to EpBmal1 caused a much weaker phenotype than the complete disruption observed in PhBmal1 knock-out animals²⁴. Alternatively, the circatidal timekeeping mechanism might differ across marine organisms, resulting from convergent or divergent evolution of a proto-circatidal clock. The only previous study at the cellular level gives some support to these possibilities: in putative circatidal cells of E. pulchra, only EpTim showed circatidal expression, whereas EpPer and EpCry2 were either arrhythmic or displayed a circadian expression pattern²⁵. In contrast, in P. hawaiensis, which lacks Tim, both PhPer and PhCry2 showed circatidal expression. Also, a recent study in which Per was knocked out in A. asahinai concluded that PER plays no role in circatidal rhythms in this insect. However, this conclusion was based on a single rhythmic animal out of 10 monitored, and no molecular rhythms were measured³⁹.

In P. hawaiensis, the loss of PhCRY2 or PhBMAL1 completely disrupted molecular and behavioral circatidal rhythms²⁴. However, in both PhPer −/− and PhClk −/− animals, residual circatidal behavioral rhythms were observed, even though molecular rhythms were severely disrupted. This demonstrates that PhPER and PhCLK are important cogs of the circatidal clock, although we surmise that residual molecular rhythmicity is present in their absence, at least in the few animals that exhibited circatidal behavior. Another bHLH/PAS transcription factor might weakly substitute for PhCLK. PhHIFα, in particular, comes to mind, since in mammals it can dimerize with BMAL1 and has been implicated in the entrainment of circadian clock by O₂ levels^40–43. Weak circatidal rhythms might be possible in the absence of functional PhPER, because repressive cryptochromes are the dominant BMAL1/CLK repressors in most species, while PER has supporting functions, such as promoting CRY nuclear entry^29,33. Our transcriptional assays indicate that this hierarchy is preserved in P. hawaiensis.

PhPer and PhCry2 mRNAs cycle in phase inside circatidal neurons, but their oscillations are antiphasic in circadian neurons²⁵ (Figure S4A to D). This suggests that the transcriptional network linking clock genes is wired differently in circadian and circatidal neurons. We therefore examined closely the impact of core clock gene mutations on PhPer and PhCry2 mRNA levels in both circadian and circatidal neurons. In both neuron types, the loss of PhPer led to moderately high mRNA levels of PhCry2, and the loss of PhCry2 to moderately high PhPer mRNA levels. This is consistent with a repressive role for both CRY2 and PER. Note that the loss of core circadian repressors does not result in peak mRNA levels in Drosophila, because interlocked feedback loops are also disrupted by the loss of core clock repressors, thus reducing DmCLK levels^44,45. PhPer and PhCry2 mRNA levels were low in PhPer −/− mutants and PhCry2 knock-outs, respectively, probably because their premature stop codons trigger nonsense-mediated mRNA decay⁴⁶.

FISH-HCR and cell culture assays thus support a standard repressive role for PhPER and PhCRY2 in both circadian and circatidal clocks. They also support the expected activator role of PhBmal1, since PhCry2 and PhPer mRNA levels are low in both clock neuron types of PhBmal1 mutants. In circadian neurons, the loss of PhClk led to low mRNA levels, supporting its canonical role as a circadian transcriptional activator along with BMAL1. Interestingly. PhCry2 levels were essentially undetectable in MP neurons. This was not linked to a technical issue, since we detected PhPer mRNAs in these circadian neurons and PhCry2 mRNAs in the DL neurons of the same brains. This suggests that PhCLK is able to work with a different partner than PhBMAL1 to weakly activate and therefore maintain constant low transcript levels of PhCry2, but is unable to do so for PhPer in circadian MP neurons. Therefore, we propose that PhPer and PhCry2 genes are activated by different sets of transcription factors in circadian neurons (Figure 7E). This could contribute to the phase difference in expression of the two repressors. Other possibilities include differences in mRNA stability, and PhCLK/PhBMAl1 controlling an intermediate positive PhCry2 regulator (Figure 7E), which would work similarly to RORα or PDP1ε in promoting antiphasic expression of mammalian Bmal1 or DmClk, respectively^6,8,9,47,48.

In circatidal neurons missing PhClk, PhPer mRNA was constantly at peak levels while PhCry2 levels were constantly low. This is unexpected, as it indicates that PhClk represses PhPer independently of PhBmal1 in circatidal neurons (Figure 7E). This is a striking deviation from its regular function in the circadian clock. It will be interesting to determine whether PhCLK represses PhPer as a homodimer, or with a novel partner, and whether its impact on PhPer expression in circatidal neurons is direct or mediated by an intermediate repressor.

In summary, transcriptional control within the core circadian and circatidal clocks is wired differently (Figure 7E). Based on the antiphasic oscillations of PhPer and PhCry2 mRNAs in circadian neurons, one would have expected the major deviation from the canonical function of core clock genes to occur in these neurons, rather than in circatidal neurons. However, the identification of a novel function for PhClk, specific to circatidal neurons, could be critical to understanding how circadian and circatidal clocks oscillate with such different periods.

Among the three historical models for circatidal rhythms, our observations, combined with those of Oliphant et al.²⁵, support the model proposed by Naylor¹⁴ of separate circatidal and circadian clocks. However, because of their overlapping mechanisms, we cannot entirely exclude the Enright model of a plastic clock¹⁵. It is possible that during development, clock neurons adopt different transcriptional wiring based on the sensory neurons they are coupled with, and thus the environmental input they receive, tidal or diurnal. In oysters, the period of core circadian gene expression changes in the presence of tides⁴⁹, and the freshwater snail Semisulcospira reiniana can favor circadian or circatidal rhythms, depending on whether it lives within or outside the tidal zone of the Kiso River^50,51. In P. hawaiensis, it will be interesting to determine what dictates the periodic fate of clock neurons, and whether this fate depends on environmental factors.

In conclusion, our results reveal that in P. hawaiensis, the evolutionary solution to the conundrum of simultaneously tracking tidal and diurnal environmental oscillations that are almost, but not exactly, harmonics, is to use the same set of core clock genes in two distinct populations of pacemaker neurons. Different autoregulatory feedback loops would set their specific periodicity. This spatial separation of the circadian and circatidal clocks likely underlies P. hawaiensis ability to integrate diurnal and tidal cues, rendering it capable of adapting its behavior and physiology to the different phase relationships of the two environmental cycles. It also likely permits plastic behavioral responses to different tidal regimens. Indeed, P. hawaiensis lives in a broad range of locations that experience different tidal patterns⁵² : semi-diurnal (the most common 12.4h tidal cycle), mixed (two tides per 24.8h, of unequal duration and amplitude) and diurnal (24.8-h tidal cycle). This work improves our understanding of the sophisticated mechanisms by which organisms cope with complex environmental periodicities. The extent to which our findings apply across marine organisms remains to be determined. Given the conservation of the circadian clock, these rules could prove universal.

Resource Availability:

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Patrick Emery [patrick.emery@umassmed.edu].

Materials availability

Plasmids and mutant animals generated in this study will be made available upon request.

Data and code availability

Original data reported in this paper will be shared by the Lead Contact upon request.
This paper does not report original codes.
Any additional information required to reanalyze the data reported in this paper will be made available by the lead contact upon request.

STAR Methods

Experimental Model and Study Participant Details

The Chicago F-strain Parhyale hawaiensis strain was used for our study. The animals were maintained in Pyrex aquariums with lids containing 30 psu artificial sea water (ASW) and ~1 cm of crushed coral substrate. The animals were reared at a temperature of 25°C under 12:12 h LD cycles with ZT0 occurring at 08:00 in the morning. Wild-type (WT) and PhBmal1 mutants²⁴ were fed baby carrots. PhCry2, PhPer and PhClk mutants were fed with a mix of ground TetraMin PLUS Tropical Flakes (Tetra, Melle, Germany), ground OSI Spirulina Flake Fish Food (Ocean Star International, Snowville, UT), ground Hikari Wheat Germ Floating Pellets for Pets (Hikari, Japan), ground Tubifex (Hagen Group, Quebec, Canada), and Kelp granules (Starwest Botanicals, Sacramento, CA). The dry food was mixed in 50 mL falcon tubes containing ASW supplemented with 50 μL of American Marine Selcon Vitamin Supplement (American Marine Inc., Ridgefield, CT) and 100 μl of KENT Marine Zoe Marine Vitamin (KENT Marine, Franklin, WI).

Methods details

CRISPR-Cas9-mediated mutagenesis

Mating pairs were isolated a day prior to embryo removal. Females were then lightly anesthetized using 0.02% clove oil in ASW. The collection of single-cell fertilized eggs was achieved as previously described²⁴. Fertilized eggs were stored in a tissue culture dish containing filtered ASW water prior to injection. Approximately 40–60 picoliters of injection mixture was injected into one-cell embryos or both cells of two-cell embryos, as previously described⁵³. The injection mixture contained 100 ng/μL or 162 ng/μL of each chemically protected guide RNA, 333 ng/μL of NLS-Cas9 protein (Synthego Co., Redwood City, CA), and 0.05% phenol red (Sigma-Aldrich). Both injection mixture concentrations generated knockouts. For each gene of interest, two guide RNAs were designed to target a functional domain or a region prior to a key functional domain. Injected embryos were then transferred to 60 mm culture dishes filled with filter-sterilized ASW and incubated at 28°C under a 12:12 h LD cycles. Embryos were transferred on a daily basis to a new culture dish containing ASW. After hatching, juvenile animals were reared individually in a custom multi-compartment insert placed inside a lidded plastic container filled with ASW. Each compartment had a mesh-covered opening that allowed water exchange with the shared bath while preventing mixing and aggression/cannibalism between individuals. Once injected animals reached sexual maturity, they were paired with other injected or wild-type individuals of the opposite sex to generate heterozygous offspring (F1) with fixed germline mutations. Heterozygotes F1 were then reared in a separate multi-compartment tank until sexual maturity. After the screening of mutations (see below), F1 males having targeted genotype were individually selected and transferred individually to a larger aquarium (20×10×10 cm) with at least 4 wild-type females. These tanks contained ~2 cm of crushed coral substrate and were fed with the food mix described above.

For the screening of mutations, adult animals were anesthetized using clove oil in ASW, after which two to three pleopods were removed and digested in 250 μL of lysis buffer (100 mM Tris, 5 mM EDTA, 200 mM NaCl, 0.2% SDS) containing 100 μg/mL of proteinase K at 50°C for 48 h. The tubes were centrifuged at 13000 rpm at 4°C for 10 min to pellet the debris. DNA was precipitated by adding 250 μL of isopropanol to the supernatant followed by centrifugation for 3 minutes at room temperature. The pellet was then washed with 500 μL of 70% ethanol, dried on a heat block at 55°Cand resuspended in 20 μL of milliQ water. PCRs were performed using Standard Taq DNA polymerase (New England BioLabs) as indicated by the manufacturer. PCR primers are listed in Table S7. The amplification program consisted of 35 cycles of denaturation at 95°C for 30 s, 30s annealing (54°C for PhCry2 primers, and 56°C for PhPer PhClk primers) and elongation at 68°C for 20 s. PCR products were run on a 3% agarose gel and the genotype was confirmed by gel purification of the products followed by sanger sequencing.

Entrainment and behavioral monitoring

Tidal entrainment was performed as previously described²⁴ with cycles of 10.3-h of high tide and 2.1-h of low tide. Low tide was simulated when water was transferred to a reservoir using a draining pump. For high tides, water was pumped back into the aquarium containing the animals. Animals were entrained to tides for a minimum period of 30 days prior to the recording of behavior. After at least five days of undisturbed entrainment (i.e. no water changes or feeding), males and females were individually loaded into polystyrene vials containing ~2 mm of crushed coral substrate and 30 psu ASW. The water height was adjusted to equal their original aquaria level, thereby maintaining a constant high tide. ParaFilm was used to close the vial and limit evaporation. Drosophila Activity Monitors (DAM) (LAM25, TriKinetics, Waltham MA) placed in a temperature and light controlled incubator (I36LL, Percival Scientific, Iowa) were used for behavioral recordings. Following a day of acclimation, swimming and roaming activity were recorded for a duration of five days with recordings conducted at one-minute intervals. After LD-only entrainment, behavior was recorded under either constant light (LL) or constant darkness conditions (DD). The behavior after tidal and LD entrainment was recorded in DD. A minimum of three sets of experiments were conducted for each condition and for each genotype.

For most behavioral studies, the mutant colonies were of mixed genotypes, containing wild-type (WT), heterozygous and homozygous mutant animals. Their genotypes were determined after recording as described above for mutant screening. For each gene, data obtained from the 2–3 different mutant lines were pooled together in later analysis as their molecular lesions were very similar and no significant difference was observed between them. For DD experiments and FISH-HCR experiments (see below), mutant colonies containing almost exclusively homozygous animals were used. These animals were nonetheless genotyped after behavioral monitoring or prior to brain dissection.

Brain sampling and processing

Animals were entrained to tides under LD cycles as described above. After LHT 12h10, five animals per sampling point were loaded separately in vials in the DAM system as described before (their behaviors are shown on figure S4E). The animals were in DD after the light was switched off at 20h00. The following day, the sampling process was initiated at 8h00. First WT animals were sampled every two hours over 24 hours (13 sampling points). For the mutants, four sampling points spaced by six hours were chosen based on WT results. The following strains were used: PhCry2 mutation 1, PhPer mutations 1 and 3 and mutation 1 for PhClk. Sampling of PhBmal1 and PhCry2 were achieved at the same time. Staining was performed in groups of two samples, for example, PhBmal1 CT0 and PhCry2 CT0. For PhPer staining, CT0 and CT6 were done simultaneously and CT12, CT18 together. The same was done for PhClk samples. A replicate of PhClk sampling was performed at CT6 and CT18 together with WT animals to eliminate the possibility of batch effects (Figure S4F). For the brain sampling, the animals were sacrificed by immersion in 4°C ASW for 20 minutes on ice. Heads were removed and placed in 4% PFA prepared in 3X PBS for 4 hours with rotation at 4°C. The heads were then washed with PBS-Tween 0.1% for 5 minutes three times. Brains were dissected in PBST using sharpened forceps. The brains were then dehydrated using a methanol dilution series (50%, 70% and 90% methanol in PBST) for five minutes each wash, followed by two brief washes in 100% methanol. Then brains were stored for a minimum of 24 hours at −20°C before staining.

HCR^™ RNA-FISH

Prior to staining, the brains were rehydrated using serial dilutions of methanol (5 min in 70%, 50% and 25%) followed by 1X10min and 2X5min washes in PBST. Half of a WT brain was added to the pooled mutant brains to ensure that no obvious technical issues occurred during the staining of the brains. They were then incubated overnight in 300 μL of detergent solution⁵⁴ (50mM Tris, 1mM EDTA, 150mM NaCl, 0.5% Tween, 1% SDS) at 4°C. The following day, the brains were pre-hybridized in 200μl hybridization buffer (Molecular Instruments Inc) at 37°C for 3 hours. The brains were then incubated in 100 μl of hybridization buffer containing 20nM PhCry2, PhPer and PhClk probes at 37°C. To determine neuronal identity, cells were co-stained using PhPer and PhElav probes (Figure S3). HCR^™ probes V3.0 were designed and synthesized by Molecular Instruments Inc, USA. After 48 h of incubation, the brains were washed 1×5min, 1×10min and 3×15min with 500μL of wash buffer (Molecular Instruments Inc) at 37°C and then 3X5min in 5XSSC supplemented by 0.1% Tween-20 at room temperature (RT). Pre-amplification consisted of 2×15min washes in 500μL of amplification buffer (Molecular Instruments Inc) at RT. Fluorescent hairpins were snap-cooled and added to 100μl of amplification buffer at a concentration of 60 nM. The brains were incubated overnight in the hairpin preparation at RT. Washes consisted of 2X5min, 2X30min and 1X5min in 500μL of 5XSSCT. The brains were then incubated in 50% glycerol in PBS containing 2μg/ml of DAPI for 1 hour at RT followed by 20 minutes in 70% glycerol. Brains were mounted in VectaShield Antifade Media (Vector Laboratories, USA) between two coverslips spaced using a Grace BioLabs SecureSeal^™ imaging spacer. Images were acquired with a LSM 900 Zeiss confocal microscope at 1μm z-stack intervals.

Transcriptional assay in HEK cells

The TK-E54 plasmid⁵⁵ harboring 3 E-Boxes driving the expression of the firefly luciferase reporter, the CMV β-galactosidase plasmid (Clontech: PT2004–5)²⁸ containing the LacZ gene under the control of the CMV promoter and the mClk⁵⁶ cDNA under the CMV promoter in pcDNA3.1 were all provided by D. Weaver (University of Massachusetts Chan Medical School). The PhBmal1 plasmid has been previously described²⁴. PhPer and PhCry2 were produced by cutting mClk plasmid using Xhol and EcoRI. gBlocks (Twist Biosciences) were designed to contain a start codon, a Flag tag (PhPer) or V5 tag (PhCry2) and the whole PhPer or PhCry2 sequences flanked by Xhol and EcoRI. Sequences used were based on Hunt et al. in silico reconstruction²⁶. PhCry2- plasmid was generated by integrating an early stop-codon into PhCry2 plasmid to mimic the mutations generated in vivo. To do so, the PhCry2 plasmid was amplified through PCR using primers designed to introduce premature stop codon. As part of this amplification, the downstream portion of the gene was removed, and a homologous overhang was included for later re-ligation of the plasmid. The resulting PhCry2 - is missing a part of the photolyase domain and the FAD binding domain, as the KO lines used in this paper. Following the amplification, Dpn1 digestion (New England Biology) was achieved following the protocol outlined by manufacturers. Following digestion, a DNA cleanup step was performed using the QIAquick PCR purification kit (Quiagen). The cleaned PhCry2- pcDNA3.1 amplicon was then religated through homologous assembly using the Gibson Assembly cloning kit (New England Biolabs) generating the finalized construct. The same protocol was used to produce the PhPer- plasmid, generating an early stop codon in the PhPer plasmid to mimic the in-vivo mutation.

HEK-293T cells were plated at 5.10⁵ cells per well in 6 well plates (Thermo Fisher Scientific) and were transfected 24-h later using Lipofectamin 2000 Transfection Reagent from Invitrogen (Waltham, MA). A total of 1600 ng of DNA was transfected per well (30 ng of TK-E54, 100 ng of PhBmal1, mClk and CMV β-gal and different increasing concentration of repressors (i.e. 0, 10, 50, 200 and 500 ng of PhCry2 and/or PhPer or PhCry2-), supplemented with empty pcDNA3.1 as needed. After transfection, plates were incubated 48 h at 37°C and 5% CO₂. They were then washed with ice-cold PBS and incubated 15 min in 350 μL of Reporter Lysis Buffer (Promega). Cell lysates were removed from the well and spun down at 13000 rpm. The supernatant was used immediately for assays or stored at −80°C.

Following the manufacturer protocol for the β-Galactosidase assay, 50 μL of the lysate was mixed with 50 μl Assay Buffer (Promega) in a 96 well plate (Corning) and incubated for 30 minutes at 37°C or until a faint yellow color appeared. The reaction was stopped using 150 μl of 1M Sodium Carbonate and the absorbance was read at 420 nm using a SpectraMax iD5 microplate reader (Molecular Devices, San Jose, CA). For the luciferase assay, 20 μl of the lysate was added to 20 μl of D-luciferin mix (0.132mg/mL D-luciferin (Sigma), 20mM Tricine, 2.67mM MgSO4, 0.1mM EDTA, 33.3mM DTT, 530uM ATP (Sigma), 270uM acetyl coenzyme A (Sigma), 265uM 4MgCO3Mg(OH)2) and the luminescence was measured using the same microplate reader. The β-Galactosidase signal was used to normalize the firefly luciferase signal.

Western Blotting

Protein amount of cell lysate was assessed using a Pierce^™ BCA Protein Assay Kit (Thermo Fisher Scientific) following the manufacturer protocol. In total, 20 μg of protein per sample was mixed with 4x Laemmli Sample buffer (Bio-Rad Laboratories) containing 2-mercaptoethanol (Sigma-Aldrich) to a final concentration of 1.05% SDS and 355 mM with 2-mercaptoethanol and boiled for 5 minutes. Samples were loaded and run at 100V on a Mini-PROTEAN TGX Stain-Free gel (Bio-Rad Laboratories). Proteins were transferred to a nitrocellulose membrane using a Trans-Blot^® SD Semi-Dry Transfer Cell (Bio-Rad Laboratories) at 15 V for 30 minutes. Membranes were washed three times 10 min in TBST and blocked in non-fat milk dissolved in TBST for an hour at 4°C. Primary antibodies were mouse anti-V5 (Invitrogen, 1:10 000), mouse anti-Flag (Sigma, 1:10 000) and mouse anti-actin (Santa Cruz biotechnology, 1:10 000). Membranes were incubated overnight in primary antibodies at 4 °C on a rocker. After washing 3 × 5 min in TBST, membranes were incubated 2h in Intercept^® (TBS) blocking buffer (LI-COR) containing IRDye 800 Goat anti-mouse (LI-COR, 1:10 000). Imaging was performed on an Odyssey DLx Imaging system (LI-COR, Lincoln, NE).

Quantification and statistical analysis

Behavior analysis

The recorded one-minute beam-crossing was averaged over 30-minute bins using DAMFileScan114 software (TriKinetics, Waltham MA). The rhythm analysis was performed with the FaasX software (https://neuropsi.cnrs.fr/en/cnn-home/francois-rouyer/faas-software/, courtesy of F. Rouyer, Centre National de la Recherche Scientifique, Gif-sur-Yvette, France)⁵⁷. Autocorrelation and χ² periodogram analysis (“power” ≥ 10 and “width” ≥ 1h) were used to determine whether an animal was rhythmic. The filter for high frequencies was on. As previously described, both “vertical swimming” and “roaming” behaviors were monitored²⁴. Animals were scored as rhythmic if vertical swimming and/or roaming were significantly rhythmic. For each animal, the period and rhythm power of the most robustly rhythmic of these two behaviors was used for subsequent analysis. For the population activity plots, behavior values were normalized for each animal by dividing each 30 minutes bin by its mean activity. Plots were generated using GraphPad Prism 10. Statistical comparisons of the power and the period were made using the same software. Outliers were identified and removed using a ROUT test with Q=1%. The normality was assessed using Shapiro-Wilk test. If the normality was respected, one-way ANOVA test was used followed by Tukey’s for multiple comparisons. Otherwise, the Kruskal-Wallis test was used, followed by Dunn’s multiple comparisons test. In instances where the comparison was between two groups, normal dataset was analyzed using Welch’s test for normal distribution. Otherwise, Mann-Whiney test was used. Statistical significance is demonstrated as ***p < 0.001; **p < 0.01; *p < 0.05. The entire dataset can be accessed in supplemental file DataS1.xlsx

As previously reported²⁴, the vast majority of wild-type animals entrained simultaneously to LD and tidal cycles showed statistically significant periodicities in the 12.4-h (“singlet”) and the harmonic 24.8-h (“doublet”) range (Table S1). After LD-only entrainment, we noticed a greater proportion of wild-type animals exhibiting a statistically significant singlet period under both LL and DD (Table S1), compared to our previous study²⁴. In other words, circadian behavior was more likely to show a bimodal daily pattern. This can also be seen on average behavior traces, particularly when behavior was monitored under DD. The percentage of rhythmic animals was also reduced under both LL and DD compared to our previous work. We are not certain of the reasons for these subtle changes in circadian behavior, but they might be related to the use of a novel entrainment chamber and new incubators for behavioral monitoring. As previously reported²⁴, under DD, the period of rhythmic behavior after entrainment to LD only was shorter than after entrainment to both LD and tidal cycles, and exposure to constant light (LL) lengthened the period of rhythmic behavior (Figure S2). This confirms that the rhythms observed after LD entrainment are of circadian nature, distinct from circatidal rhythms, even though animals were more likely to exhibit a bimodal pattern than in our previous work²⁴.

Image analysis

Image analysis was conducted using Fiji software⁵⁸. Prior to quantification, the images were smoothed and each cell group was isolated. The number of FISH dots, which reflect the expression of the gene, was counted using the RS-FISH ImageJ plugin⁵⁹. The normality of the dataset was assessed using a Shapiro-Wilk test. If a normal distribution was observed, a one-way ANOVA was used to determine if significant variations occurred though time, otherwise a Kruskal-Wallis test was used. When a statistically significant effect was observed, rhythm analyses were conducted in R (v.4.4.1)⁶⁰ using JTK CYCLE in the package MetaCycle⁶¹ for 24 h periods. P-values were adjusted using the Bonferroni method. For 12-h rhythms, the statistical difference between peaks and trough were calculated using post-hoc tests.

Supplementary Material

Data S1. Behavioral dataset and analysis. Related to Figures 1–6, Figures S2, S5, S7 and Tables S1, S2.

Individual raw swimming and roaming activity and rhythm analysis for A) PhCry2 mutant populations entrained to light dark (LD) cycles and monitored in constant light (LL). B) PhCry2 mutant populations entrained to LD cycles and monitored in constant darkness (DD). C) Wild type animals entrained to LD cycles and monitored in DD. D) PhCry2 mutant populations entrained to LD and tidal cycles with the last high tide at 13:20 and monitored in DD and constant high tide. E) PhPer mutant populations entrained to LD cycles and monitored in LL. F) PhPer mutant populations entrained to LD cycles and monitored in DD. G) PhPer mutant populations entrained to LD and tidal cycles with the last high tide at 13:20 and monitored in DD and constant high tide. H) PhPer mutant populations entrained to LD and tidal cycles with the last high tide at 19:20 and monitored in DD and constant high tide. I) PhClk mutant populations entrained to LD cycles and monitored in LL. J) PhClk mutant populations entrained to LD cycles and monitored in DD. K) PhClk mutant populations entrained to LD and tidal cycles with the last high tide at 13:20 and monitored in DD and constant high tide. L) PhClk mutant populations entrained to LD and tidal cycles with the last high tide at 19:20 and monitored in DD and constant high tide.

NIHMS2208805-supplement-1.xlsx^{(3.3MB, xlsx)}

NIHMS2208805-supplement-2.pdf^{(3.2MB, pdf)}

Document S1. Figures S1-S7 and Tables S1-S7.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Mouse anti-V5	Invitrogen	Cat#R96025
Mouse anti-Flag	Sigma Aldrich	Cat# F3165
Mouse anti-Actin (SPM161)	Santa Cruz biotechnology	Cat# Sc-56459
IRDye 800 Goat anti-mouse	LI-COR	P/N 926-32210
Chemicals, peptides, and recombinant proteins
Sea salt	Instant Ocean Sea Salt	N/A
Clove oil (Syzgium aromaticum)	Plant Therapy Essential Oil Co.	N/A
NLS-Cas9 protein	Synthego Co.	#R20SPCAS9HC
Phenol Red	Sigma-Aldrich	N/A
Proteinase K	New England Biolabs	Cat#P8107S
Standard Taq DNA Polymerase	New England Biolabs	NEB#M0273
HCR^™ Probe Hybridization Buffer (v3.0)	Molecular instrument Inc.	LOT: BPH0325
HCR^™ Probe Wash buffer V3 (v3.0)	Molecular instrument Inc.	LOT: BPM02226
HCR^™ Amplification buffer V3 (v3.0)	Molecular instrument Inc.	LOT: BAM02426
VectaShield Antifade Media	Vector Laboratories	Cat#H-1000-10
Dnp1	New England Biolabs	Cat#R0176
Lipofectamin 2000 Transfection Reagent	Invitrogen	Cat#11668027
Reporter Lysis 5X Buffer	Promega	Cat#E4030
D-luciferin	Sigma Aldrich	Cat#L9504
4x Laemmli Sample buffer	Bio-Rad Laboratories	Cat#1610747
Intercept^® (TBS) blocking buffer	LI-COR	P/N 927-60001
Mini-PROTEAN TGX Stain-free gel	Bio-Rad Laboratories	Cat#4568026
Critical commercial assays
QIAquick PCR purification kit	Quiagen	Cat#28104
Gibson Assembly Cloning kit	New England Biolabs	Cat#E5510S
β-galactosidase Enzyme Assay System	Promega	Cat#E2000
Pierce^™ BCA Protein Assay Kits	Thermo Fisher Scientific	Cat#23225
Experimental models: Cell lines
Human HEK-293T	Stephanie Puig, UMass Chan Medical School	N/A
Experimental models: Organisms/strains
Parhyale hawaiensis, Chicago-F strain	Nipam Patel, Marine Biological Laboratory (UChicago)	N/A
Parhyale hawaiensis, PhBmal −/−	Kwiatkowski et al., 2023²⁴	N/A
Parhyale hawaiensis PhCry2 −/− mutation 1	This paper	N/A
Parhyale hawaiensis PhCry2 −/− mutation 2	This paper	N/A
Parhyale hawaiensis PhPer −/− mutation 1	This paper	N/A
Parhyale hawaiensis PhPer −/− mutation 2	This paper	N/A
Parhyale hawaiensis PhPer −/− mutation 3	This paper	N/A
Parhyale hawaiensis PhClock −/− mutation 1	This paper	N/A
Parhyale hawaiensis PhClock −/− mutation 2	This paper	N/A
Oligonucleotides
PhCry2 guide RNA #1 (chemically protected) GGGCACCAAGCACCUAGAAGGU	Synthego Co.	N/A
PhCry2 guide RNA #2 (chemically protected) GGGACGAGACUCCCUGAGGUAGU	Synthego Co.	N/A
PhPer guide RNA #1 (chemically protected) GGUGGCCUAGUGGGGUGUGAGUA	Synthego Co.	N/A
PhPer guide RNA #2 (chemically protected) AGCAUACGCCAUCGCUCGUGGCC	Synthego Co.	N/A
PhClock guide RNA #1 (chemically protected) GGCGGGAGCAGUGGGGGCUCCCC	Synthego Co.	N/A
PhClock guide RNA #2 (chemically protected) GGUGGUAGCGGAAGAAGUUUGUC	Synthego Co.	N/A
Primer sets used for genotyping	See Table S7	N/A
B4-PhClock HCR^™ probe (v3.0)	Molecular instrument Inc., this paper	LOT: RTT091
B2-PhPer HCR^™ probe (v3.0)	Molecular instrument Inc., this paper	LOT: RTT709
B3-PhCry2 HCR^™ probe (v3.0)	Molecular instrument Inc., this paper	LOT: RTT817
HCR^™ Amplifier B4-647	Molecular instrument Inc.	LOT: S076626 and S072326
HCR^™ Amplifier B2-546	Molecular instrument Inc.	LOT: S049227 and S083726
HCR^™ Amplifier B3-488	Molecular instrument Inc.	LOT: S073626 and S073726
Recombinant DNA
Plasmid: TK-E54	Honma et al., 2002⁵⁵	N/A
Plasmid: CMV β-galactosidase	Clontech	PT2004-5
Plasmid: mouse Clock (mClk)	Gekakis et al., 1998⁵⁶	N/A
Plasmid: HA-PhBmal1+	Kwiatkoski et al., 2023²⁴	N/A
Plasmid: Flag-PhPer+	This paper	N/A
Plasmid: Flag-PhPer −/−	This paper	N/A
Plasmid: V5-PhCry2+	This paper	N/A
Plasmid: V5-PhCry2 −/−	This paper	N/A
Plasmid: pcDNA3.1+	Thermo Fisher Scientific	Cat#V79020
Software and algorithms
DAMFileScan114	TriKinetics	N/A
FaasX	F. Rouyer, Centre National de la Recherche Scientifique	https://neuropsi.cnrs.fr/en/cnn-home/francois-rouyer/faas-software/
GraphPad Prism 10	GraphPad	N/A
FIJI ImageJ	Schindelin et al., 2012⁵⁸	https://imagej.net/software/fiji
RS-FISH ImageJ plugin	Bahry et al., 2022⁵⁹	https://github.com/PreibischLab/RS-FISH
R	R Core Team, 2020⁶⁰	https://www.r-project.org/
MetaCycle R package	Wu et al., 2016⁶¹	https://github.com/gangwug/MetaCycle
Other
TetraMin PLUS Tropical Flakes	Tetra	N/A
OSI Spirulina Flake Fish Food	Ocean Star International	N/A
Hikari Wheat Germ Floating Pellets	Hikari	N/A
Kelp granule	Starwest Botanicals	N/A
American Marine Selcon Vitamin Supplement	American Marine Inc.	N/A
KENT Marine Zoe Marine Vitamin	KENT Marine	N/A
Drosophila Activity Monitor	TriKinetics	LAM25
Percival Incubator	Percival Scientific	I36LL
SpectraMax iD5 microplate reader	Molecular Devices	N/A
Trans-Blot SD semi-Dry Transfer Cell	Bio-Rad Laboratories	N/A
Odyssey DLx imaging system	LI-COR	N/A

Open in a new tab

Highlights:

The four core circadian clock genes sustain circatidal behavior in P. hawaiensis
These genes are essential for circatidal per and cry2 mRNA rhythms in the brain
Transcriptional wiring of clock genes differs in circadian and circatidal neurons
PhCLK represses PhPer expression independently of PhBMAL1 in circatidal neurons

Acknowledgments:

We thank Erica Kwiatkowski for generating the pAc-PhPer and pAc-PhCry2 plasmids, and for designing gRNAs for mutagenesis. We are grateful to Dr. Nipam Patel for providing advice on animal husbandry and access to his injection setup and to storage space for mutagenized colonies. We also thank Lauren North for help with animal husbandry and genotyping. This work was supported by grants from the National Science Foundation (NSF 2130765) to P.E and J.J.C.R and from the National Institute of General Medical Sciences (R35 GM145253) to P.E.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Declaration of Interests

The authors declare no competing interests.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Data S1. Behavioral dataset and analysis. Related to Figures 1–6, Figures S2, S5, S7 and Tables S1, S2.

NIHMS2208805-supplement-1.xlsx^{(3.3MB, xlsx)}

NIHMS2208805-supplement-2.pdf^{(3.2MB, pdf)}

Data Availability Statement

Original data reported in this paper will be shared by the Lead Contact upon request.
This paper does not report original codes.
Any additional information required to reanalyze the data reported in this paper will be made available by the lead contact upon request.

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[R3] 3.Dunlap JC, Loros JJ, and DeCoursey P. (2004). Chronobiology: Biological timekeeping. Sinauer Associate. [Google Scholar]

[R4] 4.Kwiatkowski ER, Rosenthal JJC, and Emery P. (2024). Clocks at sea: the genome-editing tide is rising. Trends in Genetics 40, 387–397. 10.1016/j.tig.2024.01.006. [DOI] [PubMed] [Google Scholar]

[R5] 5.Weaver DR, and Emery P. (2012). Circadian Timekeeping. In Fundamental Neuroscience, Squire Larry R., Berg Darwin, Bloom Floyd E., du Lac Sascha, Ghosh Anirvan, and Spitzer Nicholas C., eds. [Google Scholar]

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PERMALINK

Core circadian clock genes control molecular and behavioral circatidal rhythms in Parhyale hawaiensis

Victoria Louis

Zachary Bellido

Adam Helfenbein

Joshua J C Rosenthal

Patrick Emery

Summary

Graphical Abstract

Etoc blurb:

Introduction

Results

Loss of PhCry2 disrupts circadian behavior

Figure 1. PhCRY2 is required to maintain circadian rhythms in P. hawaiensis.

Loss of PhCry2 disrupts circadian molecular rhythms

PhCry2 is an essential element of the circatidal clock

Figure 2. PhCRY2 is required to maintain circatidal rhythms in P. hawaiensis.

Loss of PhPer disrupts circadian rhythms

Figure 3. PhPER is required to maintain circadian rhythms in P. hawaiensis.

PhPer sustains circatidal behavior as part of the circatidal clock

Figure 4. PhPER is required to maintain circatidal rhythms in P. hawaiensis.

PhClk is also required for circadian rhythms

Figure 5. PhCLK is required to maintain circadian rhythms in P. hawaiensis.

PhClk sustains circatidal behavior as part of the circatidal clock

Figure 6. PhCLK is required to maintain circatidal rhythms in P. hawaiensis.

PhBmal1 also controls circadian and circatidal expression, but PhBMAl1 and PhCLK functions differ in circatidal neurons

Figure 7. PhBMAL1 is part of the molecular circadian and circatidal clocks in Parhyale.

Discussion

Resource Availability:

Lead contact

Materials availability

Data and code availability

STAR Methods

Experimental Model and Study Participant Details

Methods details

CRISPR-Cas9-mediated mutagenesis

Entrainment and behavioral monitoring

Brain sampling and processing

HCR™ RNA-FISH

Transcriptional assay in HEK cells

Western Blotting

Quantification and statistical analysis

Behavior analysis

Image analysis

Supplementary Material

Highlights:

Acknowledgments:

Footnotes

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

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HCR^™ RNA-FISH